Which cryogenic vials and caps are ideal for storing glycerol stocks?

Which cryogenic vials and caps are ideal for storing glycerol stocks?

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There are internally threaded, externally threaded, natural caps. For the vials, there are skirts, star-footed, and various bottoms. Why do any of these things matter?

For any kind of frozen cell stock (for cell culture or bacteria) we routinely use freezing vials similar to these:

These tubes are internally threaded with a rubber grommet to seal the lid and make the tube air and water tight. I suspect the air tight seal prevents condensation from entering the tube and freezing within the tube, but I've never had a problem with either choice of tube.

The tubes don't really matter at -80 C for a glycerol stab. Freezing cells in liquid nitrogen on the other hand has a huge impact when storing in liquid nitrogen. Some tubes are meant to be stored in the liquid versus the vapour phase of nitrogen and not matching the tube to it's intended storage condition can result in explosion of the tube.

Freezing Cells

Cell lines in continuous culture are likely to suffer undesirable outcomes such as genetic drift, senescence, and microbial contamination, and even the best-run laboratories can experience equipment failure. An established cell line is a valuable resource, and its replacement is expensive and time consuming. Therefore, it is vitally important that they are frozen down and preserved for long-term storage. A properly maintained frozen cell stock is an important part of cell culture.

As soon as a small surplus of cells becomes available from subculturing, the best preservative method is to keep them frozen as a seed stock, protected, and not be made available for general laboratory use. Working stocks can be prepared and replenished from frozen seed stocks. If the seed stocks become depleted, cryopreserved working stocks can then serve as a source for preparing a fresh seed stock with a minimum increase in generation number from the initial freezing.

The general freezing method is the same for adherent and suspension cells, except that adherent cells need to be removed from the culture plates before starting the freezing procedure. The best method for cryopreserving cultured cells is storing them in liquid nitrogen in complete medium in the presence of a cryoprotective agent such as dimethyl sulfoxide (DMSO). Cryoprotective agents reduce the freezing point of the medium and allow a slower cooling rate, greatly reducing the risk of ice crystal formation, which can damage cells and cause cell death.

Note: A DMSO solution is known to facilitate the entry process of organic molecules into tissues. Handle reagents containing DMSO using equipment and practices appropriate for the hazards posed by such materials. Dispose of the reagents in compliance with local regulations.

Freezing medium
Always use the recommended freezing medium for cryopreserving your cells. The freezing medium should contain a cryoprotective agent such as DMSO or glycerol. You may also use a specially formulated complete cryopreservation medium such as Gibco Recovery Cell Culture Freezing Medium or Gibco Synth-a-Freeze Cryopreservation Medium.

    is a ready-to-use complete cryopreservation medium for mammalian cell cultures, containing an optimized ratio of fetal bovine serum to bovine serum for improved cell viability and cell recovery after thawing. is a chemically defined, protein-free, sterile cryopreservation medium containing 10% DMSO that is suitable for the cryopreservation of many stem and primary cell types with the exception of melanocytes.

Which cryogenic vials and caps are ideal for storing glycerol stocks? - Biology

  • Effective maintenance of stock cultures is essential for QC, method validation and research purposes
  • Repeated subculturing may eventually lead to contamination, loss of viability and genotypic/phenotypic changes
  • Freeze-drying and cryogenic storage are preferred, but may not be practical for smaller laboratories
  • Cryoprotectant beads allow routine labs to maintain a stock culture collection simply and at low cost.

One of the most important, yet often neglected, tasks in any routine microbiology laboratory is to maintain a collection of bacterial and fungal stock cultures. In a busy laboratory it is all too easy for the stock culture collection to deteriorate into a jumble of poorly labelled, partially dried-out agar slant cultures at the back of a refrigerator. But it does not have to be, nor should it be, like that.

There are a number of reasons why a microbiology laboratory needs stock cultures in good condition. The typical stock culture collection may contain isolates that fall into one or more of the following categories:

  • Reference strains for quality control of culture media and methods
  • Isolates used in the preparation of inoculated samples and specimens for quality control and training purposes
  • Reference strains for the development and validation of new methods
  • Pathogens and spoilage organisms lated during routine testing or in the investigation of contamination problems
  • Cultures used in microbiological assays
  • Isolates required for research purposes

It is difficult to conceive of a laboratory that does not need stock cultures for at least one of these reasons, even if it is simply a matter of keeping a few reference strains for QC purposes.

Methods for Establishing and Maintaining a Culture Collection

  • Risk of contamination – in time, it is possible for important isolates to be completely overgrown by contaminants
  • Loss of viability – if subculturing is not carried out at the required intervals and the cultures are inadequately stored, sensitive isolates may lose viability and be irrecoverable
  • Continued growth at chill temperatures – some organisms, such as Listeria monocytogenes, are capable of slow growth at 0 o C or even less
  • Labelling mistakes – subculturing a large number of agar slants many times introduces a significant chance of a culture being wrongly labelled
  • Genetic drift and mutation – every subculture carries a potential for genotypic and phenotypic changes, such as loss of virulence and resistance factors, or reduced motility, to occur

From this it is clear that unless stock cultures are scrupulously maintained they can cause serious problems in the laboratory and even a well maintained collection may give misleading results in time. In fact, the American Type Culture Collection (ATCC) recommends that no more than five passages (subcultures) should be made from the original type strain, with the first passage being defined as the culture prepared from the vial supplied by ATCC.

Alternative methods Fortunately, laboratories are not limited to the traditional agar slant stock culture these days, and there are other methods available to make the process of maintaining the collection easier and to overcome the problems described above.

Freeze-drying Also known as lyophilisation, freeze-drying is a method that can be used to suspend the metabolism of bacterial and fungal cultures and to stabilise them for long-term storage.

A thick suspension of bacterial cells or fungal spores is first prepared in a suitable suspending medium, such as 10% skim milk, or a specific lyophilisation buffer. This suspension is then dispensed into small glass vials and frozen. Once frozen, the plugged or loosely capped vials are placed in the drying chamber of a freeze dryer and dried under vacuum for 2 – 24 hours to remove water in the frozen state. A secondary drying step may also be applied by attaching the vials to the manifold of the freeze-dryer for a further 2 – 12 hours. When drying is complete the vial is sealed and then stored in the dark at 8 o C or less.

Many bacterial and fungal species will remain stable and viable for at least a year under these conditions and in some cases cultures have been successfully resuscitated many years later. However, some fastidious or delicate species may be damaged by the process, or may not remain viable. For example, the cells of Campylobacter spp and moulds that produce large fragile spores require careful handling as they are very susceptible to damage and can only be stored for shorter periods.

Freeze-drying is the preferred method keeping reference strains and other isolates for long periods and is standard practice for large commercial and national culture collections. Many culture collections supply freeze-dried cultures to laboratories in glass vials, but it is also possible to obtain some QC strains in easy-to-open containers that include hydration media and an inoculating swab to simplify resuscitation and subculture.

Freeze-drying is probably not suitable for smaller laboratories that only need to maintain a small stock culture collection. As well as from the need for costly freeze-drying equipment, the process is time consuming and requires skilled staff to ensure that no contamination occurs.

Cryogenic Storage An alternative method of storing cultures under conditions that suspend metabolism is so-called cryogenic storage, usually using liquid nitrogen. Suspensions of bacterial cells or fungal spores are prepared in a cryoprotectant medium, generally containing 10-15% glycerol to minimise damage during freezing. The suspension is then dispensed into suitable containers, such as small screw-capped vials, which are then immersed in, or suspended above, liquid nitrogen.

The temperature of liquid nitrogen is -196 o C, well below the temperature at which all metabolic activity is thought to cease. As with freeze-drying, not all microbial cultures will survive the process, but those that do may survive for many years. Indeed viability for some isolates may be better than for freeze-dried cultures. However, cryogenic storage is expensive, requires significant amounts of liquid nitrogen and is probably only suitable for larger reference laboratories with extensive culture collections.

Frozen storage on cryoprotectant beads Perhaps the most practical method of long term microbial stock culture storage for smaller laboratories and those engaged in routine testing is to freeze cultures onto porous beads designed to allow cells and spores to attach to their surface – providing some degree of protection against damage during freezing.

There are a number of readily available commercial products that laboratories can use to do this, but all operate in much the same way.

Outline method 1. Prepare a suspension of cells or spores of the isolate to be stored, preferably using an 18 to 24 hour old culture grown on solid medium. The colonial growth is suspended in a cryoprotectant fluid, such as Brucella Broth with Glycerol, in a cryogenic vial containing 20-30 of the porous beads.

2. The suspension is mixed by inversion and then allowed to stand at room temperature for 15-20 minutes. During this period a layer of cells will adhere to the porous beads, which have a large surface area to volume ratio.

3. The excess cryoprotectant liquid is then removed by aspiration and the vial is immediately transferred to a freezer, preferably at -70 o C or less (although the beads can be kept at -20 o C, they will lose viability more quickly than at lower temperatures). The cryogenic vials may be kept in aluminium ‘cryoblocs’ to help prevent rapid thawing when they are removed from the freezer.

4. To resuscitate a culture, remove a single bead quickly and use it to inoculate a solid, non-selective medium. The vial should be resealed and returned to the freezer immediately so that the remainder of the culture is not allowed to thaw.

This method allows stable stock cultures to be kept for extended periods, providing that the temperature is kept at -70 o C or less. The cultures are also less vulnerable to contamination. No specialised, or expensive equipment is required and so the method is suitable for small routine laboratories. The use of porous beads also allows laboratories to make better use of freeze-dried reference cultures obtained from large collections and reduces the need for frequent replacement. Other considerations Correct labelling is a vital part of maintaining any stock culture collection. Cryogenic vials are usually designed for labelling with permanent marker pens and are also available with colour-coded tops to help laboratories identify particular categories of cultures.

Accredited laboratories will normally need to keep records of the maintenance of their stock cultures where they are used for QC and validation purposes. It may help to prepare specific documentation for this, but basic computer spreadsheets and more specialised laboratory software applications can also be used. It is important to log every subculturing of all stock cultures so that the collection can be maintained in good condition and replaced when necessary.

Outsourcing For laboratories with extensive stock culture collections, perhaps including unique strains isolated from patients, the environment, or spoiled products, it may be helpful to allow a specialist reference laboratory to help maintain the collection. Several commercial culture collections offer specialist freeze-drying services and will also store important cultures for clients. This may be vital if the integrity of the laboratory collection is compromised by an event such as a faulty freezer.

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Water-bears (Tardigrada), microscopic multicellular organisms, can survive freezing by replacing most of their internal water with the sugar trehalose, preventing it from crystallization that otherwise damages cell membranes. Mixtures of solutes can achieve similar effects. Some solutes, including salts, have the disadvantage that they may be toxic at intense concentrations. In addition to the water-bear, wood frogs can tolerate the freezing of their blood and other tissues. Urea is accumulated in tissues in preparation for overwintering, and liver glycogen is converted in large quantities to glucose in response to internal ice formation. Both urea and glucose act as "cryoprotectants" to limit the amount of ice that forms and to reduce osmotic shrinkage of cells. Frogs can survive many freeze/thaw events during winter if no more than about 65% of the total body water freezes. Research exploring the phenomenon of "freezing frogs" has been performed primarily by the Canadian researcher, Dr. Kenneth B. Storey. [ citation needed ]

Freeze tolerance, in which organisms survive the winter by freezing solid and ceasing life functions, is known in a few vertebrates: five species of frogs (Rana sylvatica, Pseudacris triseriata, Hyla crucifer, Hyla versicolor, Hyla chrysoscelis), one of salamanders (Salamandrella keyserlingii), one of snakes (Thamnophis sirtalis) and three of turtles (Chrysemys picta, Terrapene carolina, Terrapene ornata). [3] Snapping turtles Chelydra serpentina and wall lizards Podarcis muralis also survive nominal freezing but it has not been established to be adaptive for overwintering. In the case of Rana sylvatica one cryopreservant is ordinary glucose, which increases in concentration by approximately 19 mmol/l when the frogs are cooled slowly. [3]

One early theoretician of cryopreservation was James Lovelock. In 1953, he suggested that damage to red blood cells during freezing was due to osmotic stress, [4] and that increasing the salt concentration in a dehydrating cell might damage it. [5] [6] In the mid-1950s, he experimented with the cryopreservation of rodents, determining that hamsters could be frozen with 60% of the water in the brain crystallized into ice with no adverse effects other organs were shown to be susceptible to damage. [7]

Cryopreservation was applied to human materials beginning in 1954 with three pregnancies resulting from the insemination of previously frozen sperm. [8] Fowl sperm was cryopreserved in 1957 by a team of scientists in the UK directed by Christopher Polge. [9] During 1963, Peter Mazur, at Oak Ridge National Laboratory in the U.S., demonstrated that lethal intracellular freezing could be avoided if cooling was slow enough to permit sufficient water to leave the cell during progressive freezing of the extracellular fluid. That rate differs between cells of differing size and water permeability: a typical cooling rate around 1 °C/minute is appropriate for many mammalian cells after treatment with cryoprotectants such as glycerol or dimethyl sulphoxide, but the rate is not a universal optimum. [10]

On April 22, 1966, the first human body was frozen—though it had been embalmed for two months—by being placed in liquid nitrogen and stored at just above freezing. The elderly woman from Los Angeles, whose name is unknown, was soon thawed out and buried by relatives. The first human body to be frozen with the hope of future revival was James Bedford's, a few hours after his cancer-caused death in 1967. [11] Bedford is the only cryonics patient frozen before 1974 still preserved today. [12]

Storage at very low temperatures is presumed to provide an indefinite longevity to cells, although the actual effective life is rather difficult to prove. Researchers experimenting with dried seeds found that there was noticeable variability of deterioration when samples were kept at different temperatures – even ultra-cold temperatures. Temperatures less than the glass transition point (Tg) of polyol's water solutions, around −136 °C (137 K −213 °F), seem to be accepted as the range where biological activity very substantially slows, and −196 °C (77 K −321 °F), the boiling point of liquid nitrogen, is the preferred temperature for storing important specimens. While refrigerators, freezers, and extra-cold freezers are used for many items, generally the ultra-cold of liquid nitrogen is required for successful preservation of the more complex biological structures to virtually stop all biological activity.

Phenomena which can cause damage to cells during cryopreservation mainly occur during the freezing stage, and include solution effects, extracellular ice formation, dehydration, and intracellular ice formation. Many of these effects can be reduced by cryoprotectants. Once the preserved material has become frozen, it is relatively safe from further damage. [13]

Solution effects As ice crystals grow in freezing water, solutes are excluded, causing them to become concentrated in the remaining liquid water. High concentrations of some solutes can be very damaging. Extracellular ice formation When tissues are cooled slowly, water migrates out of cells and ice forms in the extracellular space. Too much extracellular ice can cause mechanical damage to the cell membrane due to crushing. Dehydration Migration of water, causing extracellular ice formation, can also cause cellular dehydration. The associated stresses on the cell can cause damage directly. Intracellular ice formation While some organisms and tissues can tolerate some extracellular ice, any appreciable intracellular ice is almost always fatal to cells.

The main techniques to prevent cryopreservation damages are a well-established combination of controlled rate and slow freezing and a newer flash-freezing process known as vitrification.

Slow programmable freezing Edit

Controlled-rate and slow freezing, also known as slow programmable freezing (SPF), [14] is a set of well established techniques developed during the early 1970s which enabled the first human embryo frozen birth Zoe Leyland during 1984. Since then, machines that freeze biological samples using programmable sequences, or controlled rates, have been used all over the world for human, animal, and cell biology – "freezing down" a sample to better preserve it for eventual thawing, before it is frozen, or cryopreserved, in liquid nitrogen. Such machines are used for freezing oocytes, skin, blood products, embryos, sperm, stem cells, and general tissue preservation in hospitals, veterinary practices and research laboratories around the world. As an example, the number of live births from frozen embryos 'slow frozen' is estimated at some 300,000 to 400,000 or 20% of the estimated 3 million in vitro fertilization (IVF) births. [15]

Lethal intracellular freezing can be avoided if cooling is slow enough to permit sufficient water to leave the cell during progressive freezing of the extracellular fluid. To minimize the growth of extracellular ice crystals and recrystallization, [16] biomaterials such as alginates, polyvinyl alcohol or chitosan can be used to impede ice crystal growth along with traditional small molecule cryoprotectants. [2] That rate differs between cells of differing size and water permeability: a typical cooling rate of about 1 °C/minute is appropriate for many mammalian cells after treatment with cryoprotectants such as glycerol or dimethyl sulfoxide, but the rate is not a universal optimum. The 1 °C / minute rate can be achieved by using devices such as a rate-controlled freezer or a benchtop portable freezing container. [17]

Several independent studies have provided evidence that frozen embryos stored using slow-freezing techniques may in some ways be 'better' than fresh in IVF. The studies indicate that using frozen embryos and eggs rather than fresh embryos and eggs reduced the risk of stillbirth and premature delivery though the exact reasons are still being explored.

Vitrification Edit

Researchers Greg Fahy and William F. Rall helped to introduce vitrification to reproductive cryopreservation in the mid-1980s. [18] As of 2000, researchers claim vitrification provides the benefits of cryopreservation without damage due to ice crystal formation. [19] The situation became more complex with the development of tissue engineering as both cells and biomaterials need to remain ice-free to preserve high cell viability and functions, integrity of constructs and structure of biomaterials. Vitrification of tissue engineered constructs was first reported by Lilia Kuleshova, [20] who also was the first scientist to achieve vitrification of oocytes, which resulted in live birth in 1999. [21] For clinical cryopreservation, vitrification usually requires the addition of cryoprotectants before cooling. Cryoprotectants are macromolecules added to the freezing medium to protect cells from the detrimental effects of intracellular ice crystal formation or from the solution effects, during the process of freezing and thawing. They permit a higher degree of cell survival during freezing, to lower the freezing point, to protect cell membrane from freeze-related injury. Cryoprotectants have high solubility, low toxicity at high concentrations, low molecular weight and the ability to interact with water via hydrogen bonding.

Instead of crystallizing, the syrupy solution becomes an amorphous ice—it vitrifies. Rather than a phase change from liquid to solid by crystallization, the amorphous state is like a "solid liquid", and the transformation is over a small temperature range described as the "glass transition" temperature.

Vitrification of water is promoted by rapid cooling, and can be achieved without cryoprotectants by an extremely rapid decrease of temperature (megakelvins per second). The rate that is required to attain glassy state in pure water was considered to be impossible until 2005. [22]

Two conditions usually required to allow vitrification are an increase of viscosity and a decrease in the freezing temperature. Many solutes do both, but larger molecules generally have a larger effect, particularly on viscosity. Rapid cooling also promotes vitrification.

For established methods of cryopreservation, the solute must penetrate the cell membrane in order to achieve increased viscosity and decrease the freezing temperature inside the cell. Sugars do not readily permeate through the membrane. Those solutes that do, such as dimethyl sulfoxide, a common cryoprotectant, are often toxic in intense concentration. One of the difficult compromises of vitrifying cryopreservation concerns limiting the damage produced by the cryoprotectant itself due to cryoprotectant toxicity. Mixtures of cryoprotectants and the use of ice blockers have enabled the Twenty-First Century Medicine company to vitrify a rabbit kidney to −135 °C with their proprietary vitrification mixture. Upon rewarming, the kidney was transplanted successfully into a rabbit, with complete functionality and viability, able to sustain the rabbit indefinitely as the sole functioning kidney. [23]

Persufflation Edit

Blood can be replaced with inert noble gases and/or metabolically vital gases like oxygen, so that organs can cool more quickly and less antifreeze is needed. Since regions of tissue are separated by gas, small expansions do not accumulate, thereby protecting against shattering. [24] A small company, Arigos Biomedical, "has already recovered pig hearts from the 120 degrees below zero", [25] although the definition of "recovered" is not clear. Pressures of 60 atm can help increase heat exchange rates. [26] Gaseous oxygen perfusion/persufflation can enhance organ preservation relative to static cold storage or hypothermic machine perfusion, since the lower viscosity of gases, may help reach more regions of preserved organs and deliver more oxygen per gram tissue. [27]

Generally, cryopreservation is easier for thin samples and suspended cells, because these can be cooled more quickly and so require lesser doses of toxic cryoprotectants. Therefore, cryopreservation of human livers and hearts for storage and transplant is still impractical.

Nevertheless, suitable combinations of cryoprotectants and regimes of cooling and rinsing during warming often allow the successful cryopreservation of biological materials, particularly cell suspensions or thin tissue samples. Examples include:

    in semen cryopreservation
    • Special cells for transfusion like platelets (Thrombosomes by Cellphire) . It is optimal in high concentration of synthetic serum, stepwise equilibration and slow cooling. [28] in a Cord blood bank

    Embryos Edit

    Cryopreservation for embryos is used for embryo storage, e.g., when in vitro fertilization (IVF) has resulted in more embryos than is currently needed.

    One pregnancy and resulting healthy birth has been reported from an embryo stored for 27 years after the successful pregnancy of an embryo from the same batch three years earlier. [30] Many studies have evaluated the children born from frozen embryos, or “frosties”. The result has been uniformly positive with no increase in birth defects or development abnormalities. [31] A study of more than 11,000 cryopreserved human embryos showed no significant effect of storage time on post-thaw survival for IVF or oocyte donation cycles, or for embryos frozen at the pronuclear or cleavage stages. [32] Additionally, the duration of storage did not have any significant effect on clinical pregnancy, miscarriage, implantation, or live birth rate, whether from IVF or oocyte donation cycles. [32] Rather, oocyte age, survival proportion, and number of transferred embryos are predictors of pregnancy outcome. [32]

    Ovarian tissue Edit

    Cryopreservation of ovarian tissue is of interest to women who want to preserve their reproductive function beyond the natural limit, or whose reproductive potential is threatened by cancer therapy, [33] for example in hematologic malignancies or breast cancer. [34] The procedure is to take a part of the ovary and perform slow freezing before storing it in liquid nitrogen whilst therapy is undertaken. Tissue can then be thawed and implanted near the fallopian, either orthotopic (on the natural location) or heterotopic (on the abdominal wall), [34] where it starts to produce new eggs, allowing normal conception to occur. [35] The ovarian tissue may also be transplanted into mice that are immunocompromised (SCID mice) to avoid graft rejection, and tissue can be harvested later when mature follicles have developed. [36]

    Oocytes Edit

    Human oocyte cryopreservation is a new technology in which a woman's eggs (oocytes) are extracted, frozen and stored. Later, when she is ready to become pregnant, the eggs can be thawed, fertilized, and transferred to the uterus as embryos. Since 1999, when the birth of the first baby from an embryo-derived from vitrified-warmed woman's eggs was reported by Kuleshova and co-workers in the journal of Human Reproduction, [20] this concept has been recognized and widespread. This breakthrough in achieving vitrification of a woman's oocytes made an important advance in our knowledge and practice of the IVF process, as the clinical pregnancy rate is four times higher after oocyte vitrification than after slow freezing. [37] Oocyte vitrification is vital for preserving fertility in young oncology patients and for individuals undergoing IVF who object, for either religious or ethical reasons, to the practice of freezing embryos.

    Semen Edit

    Semen can be used successfully almost indefinitely after cryopreservation. The longest reported successful storage is 22 years. [38] It can be used for sperm donation where the recipient wants the treatment in a different time or place or as a means of preserving fertility for men undergoing vasectomy or treatments that may compromise their fertility, such as chemotherapy, radiation therapy or surgery.

    Testicular tissue Edit

    Cryopreservation of immature testicular tissue is a developing method to avail reproduction to young boys who need to have gonadotoxic therapy. Animal data are promising since healthy offspring have been obtained after transplantation of frozen testicular cell suspensions or tissue pieces. However, none of the fertility restoration options from frozen tissue, i.e. cell suspension transplantation, tissue grafting and in vitro maturation (IVM) has proved efficient and safe in humans as yet. [39]

    Moss Edit

    Cryopreservation of whole moss plants, especially Physcomitrella patens, has been developed by Ralf Reski and coworkers [40] and is performed at the International Moss Stock Center. This biobank collects, preserves, and distributes moss mutants and moss ecotypes. [41]

    Mesenchymal stromal cells (MSCs) Edit

    MSCs, when transfused immediately within a few hours post-thawing, may show reduced function or show decreased efficacy in treating diseases as compared to those MSCs which are in log phase of cell growth (fresh). As a result, cryopreserved MSCs should be brought back into the log phase of cell growth in in vitro culture before these are administered for clinical trials or experimental therapies. Re-culturing of MSCs will help in recovering from the shock the cells get during freezing and thawing. Various clinical trials on MSCs have failed which used cryopreserved products immediately post-thaw as compared to those clinical trials which used fresh MSCs. [42]

    Bacteria and fungi can be kept short-term (months to about a year, depending) refrigerated, however, cell division and metabolism is not completely arrested and thus is not an optimal option for long-term storage (years) or to preserve cultures genetically or phenotypically, as cell divisions can lead to mutations or sub-culturing can cause phenotypic changes. A preferred option, species-dependent, is cryopreservation. Nematode worms are the only multicellular eukaryotes that have been shown to survive cryopreservation. [43] Shatilovich AV, Tchesunov AV, Neretina TV, Grabarnik IP, Gubin SV, Vishnivetskaya TA, Onstott TC, Rivkina EM (May 2018). "Viable Nematodes from Late Pleistocene Permafrost of the Kolyma River Lowland". Doklady Biological Sciences. 480 (1): 100–102. doi:10.1134/S0012496618030079. PMID 30009350. S2CID 49743808.

    Fungi Edit

    Fungi, notably zygomycetes, ascomycetes, and higher basidiomycetes, regardless of sporulation, are able to be stored in liquid nitrogen or deep-frozen. Cryopreservation is a hallmark method for fungi that do not sporulate (otherwise other preservation methods for spores can be used at lower costs and ease), sporulate but have delicate spores (large or freeze-dry sensitive), are pathogenic (dangerous to keep metabolically active fungus) or are to be used for genetic stocks (ideally to have an identical composition as the original deposit). As with many other organisms, cryoprotectants like DMSO or glycerol (e.g. filamentous fungi 10% glycerol or yeast 20% glycerol) are used. Differences between choosing cryoprotectants are species (or class) dependent, but generally for fungi penetrating cryoprotectants like DMSO, glycerol or polyethylene glycol are most effective (other non-penetrating ones include sugars mannitol, sorbitol, dextran, etc.). Freeze-thaw repetition is not recommended as it can decrease viability. Back-up deep-freezers or liquid nitrogen storage sites are recommended. Multiple protocols for freezing are summarized below (each uses screw-cap polypropylene cryotubes): [44]

    Bacteria Edit

    Many common culturable laboratory strains are deep-frozen to preserve genetically and phenotypically stable, long-term stocks. [45] Sub-culturing and prolonged refrigerated samples may lead to loss of plasmid(s) or mutations. Common final glycerol percentages are 15, 20, and 25. From a fresh culture plate, one single colony of interest is chosen and liquid culture is made. From the liquid culture, the medium is directly mixed with an equal amount of glycerol the colony should be checked for any defects like mutations. All antibiotics should be washed from the culture before long-term storage. Methods vary, but mixing can be done gently by inversion or rapidly by vortex and cooling can vary by either placing the cryotube directly at −50 to −95 °C, shock-freezing in liquid nitrogen or gradually cooling and then storing at −80 °C or cooler (liquid nitrogen or liquid nitrogen vapor). Recovery of bacteria can also vary, namely, if beads are stored within the tube then the few beads can be used to plate or the frozen stock can be scraped with a loop and then plated, however, since only little stock is needed the entire tube should never be completely thawed and repeated freeze-thaw should be avoided. 100% recovery is not feasible regardless of methodology. [46] [47] [48]

    Worms Edit

    The microscopic soil-dwelling nematode roundworms Panagrolaimus detritophagus and Plectus parvus are the only eukaryotic organisms that have been proven to be viable after long-term cryopreservation to date. In this case, the preservation was natural rather than artificial, due to permafrost.

    Vertebrates Edit

    Several animal species, including fish, amphibians, and reptiles have been shown to tolerate freezing. These species include at least four species of frogs (Pseudacris crucifer, Hyla versicolor, Pseudacris triseriata, Lithobates sylvaticus) and several species of turtles (Terrapene carolina, hatchling Chrysemys picta), lizards, and snakes are freeze tolerant and have developed adaptations for surviving freezing. While some frogs hibernate underground or in water, body temperatures still drop to −5 to −7 °C, causing them to freeze. The Wood frog (Lithobates sylvaticus) can withstand repeated freezing, during which about 65% of its extracellular fluid is converted to ice. [45]

    Preserving Microorganisms Using Microencapsulation?

    Microencapsulation, where cells are entrapped in a matrix prior to storage, has been proposed as a long term microbial preservation method which does not expose the microorganisms to the harsh stresses of freezing and drying. The matrix shields the cells and increases stability during storage. Microencapsulation of probiotic bacteria in calcium alginate has been shown to improve their viability when stored at -80°C. Electrospinning and electrospraying, where microorganisms are trapped in nanofibers and droplets respectively, have also been used to preserve the viability of sensitive probiotic bacteria [6].

    Microbial preservation techniques enable the viability of microorganisms to be maintained, paving the way for their potential to be explored for years to come. With the right methods, some of these microorganisms may still be around in freezers long after we are gone, waiting to be revived by future generations of researchers!


    Using Koch’s postulates for the identification of pathogenic microbes, Ogston identified the etiological agent of suppurative abscesses (Ogston, 1883). The name Staphylococcus aureus was chosen to distinguish this species with its characteristic yellow colony pigment from another staphylococcal commensal that forms white colonies (Staphylococcus albus, now designated Staphylococcus epidermidis) (Rosenbach, 1884 Götz et al., 2006). S. aureus displays several striking microbiological properties, e.g., the microbe binds immunoglobulins and agglutinates with or coagulates blood and plasma (Loeb, 1903 Much, 1908 Forsgren and Sjöquist, 1966 Cheng et al., 2011). These traits have been useful for the early and rapid diagnosis of S. aureus infections (for a historical account of the coagulase test, follow the link

    All Staphylococci grow in clusters, a feature that can be visualized by microscopy and accounts for the Greek name σταΦυλoκoκκoς or grape-like berry. Clustering is caused by the incomplete separation of daughter cells following division in three alternating perpendicular planes (Tzagoloff and Novick, 1977 Giesbrecht et al., 1998). S. aureus cells appear perfectly spherical with a diameter of

    1 μm (Giesbrecht et al., 1998).S. aureus also produces catalase when applied to colony material, the catalase test is a rapid, useful test to distinguish staphylococci from other Gram-positive bacteria such as streptococci.

    S. aureus is a facultative anaerobe that grows by aerobic respiration or by fermentation, which yields principally lactic acid. The bacterium metabolizes glucose via the pentose phosphate pathway (Reizer et al., 1998). There is no evidence for the existence of the Entner-Doudoroff pathway however, enzymes of the entire tricarboxylic acid cycle and a typical F0F1-ATPase are encoded by the genome of S. aureus (Kuroda et al., 2001). Upon glucose depletion, S. aureus cells growing in aerobic conditions oxidize D-galactose, acetate, succinate and malate. An excellent summary of these metabolic pathways was recently published (Götz et al., 2006).

    CAUTION: S. aureus is a highly virulent and adaptable pathogen with the ability to infect, invade, persist, and replicate in any human tissue including skin, bone, visceral organs, or vasculature (Lowy, 1998). The organism has been placed in Risk Group Level 2. All manipulations with S. aureus strains must be performed following biosafety level 2 measures including experimental work in certified biosafety cabinets. Guidelines for BSL2 practice can be obtained from the latest edition of Biosafety in Microbiological and Biomedical Laboratories (BMBL, 5th Edition) via the following CDC Web link:




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    Representative Results

    An example of C. difficile grown on BHIS and Columbia anaerobic sheep blood agar media can be seen in Figure 2. C. difficile forms irregular colonies that are flat and possess a ground-glass appearance which is evident on both media. Here, an erythromycin-sensitive clinical isolate of C. difficile, 630E 30 , is grown on BHIS agar, an enriched, non-selective medium, for 24 hr at 37 ଌ (Figure 2A). Colonies on Columbia anaerobic sheep blood agar appear similar to those grown on BHIS under white light (Figure 2B) however, the use of this medium also provides for detection of the greenish or chartreuse fluorescence exhibited by C. difficile under long-wave ultraviolet (UV) light 15 (Figure 2C). C. difficile colonies on TCCFA agar look similar to growth on BHIS agar. Because of the presence of two antibiotics in TCCFA medium, a time period of 48 hr growth is necessary before enumerating colonies.

    Figure 1. The Coy Laboratories Type C vinyl chamber and its components. (A) A Coy Laboratories Type C vinyl chamber which provides workspace for a single individual at one time (42 in. x 32 in.). It contains a catalyst fan box (back left corner) which circulates and heats the air, and holds the Stak-Pak containing the palladium catalyst required to reduce any oxygen contamination. (B) The airlock serves as an interchange and provides a mechanism for the transfer of materials in and out of the chamber while preventing significant oxygen contamination within the anaerobic environment. The airlock has two doors: one providing access to the outside of the airlock and the other providing access to the interior of the anaerobic chamber. The airlock is programmable and allows for customized cycles for entry into the chamber. It is operable in automatic, semi-manual and manual modes. (C) Attached, flexible latex gloves are provided which allow full range of motion and reach within the chamber. The gloves are secured to a specialized cuff attached to the vinyl sleeves with vinyl adhesive, permitting replacement of gloves without disrupting the anaerobic atmosphere of the chamber. Neoprene gloves are also available. (D) The Model 10 Gas Analyzer continuously monitors both oxygen and hydrogen levels providing an instantaneous readout of the atmosphere within the chamber. This unit allows for immediate alerts if a leak occurs, an incorrect gas mix is used or additional problems arise via audible alarms and a flashing LED light.

    Figure 2. The appearance of C. difficile colonies on various media. The characteristic flat, irregular, ground-glass appearance is evident with an erythromycin-sensitive clinical isolate of C. difficile, 630E 30 , grown on BHIS agar for 24 hr (A) and Columbia anaerobic sheep blood agar for 48 hr under white light (B) and long wave ultraviolet light (C).

    What type of cryovial to use to preserve samples at -80C?

    So I'm processing 700 individual soil samples into a -80C freezer with the hope to do environmental DNA extraction on each sample using soil DNA extraction kits, about a month or two down the line.

    I've already stored many of the soil samples into the -80C freezer within this exact type of Cryovial (externally threaded)

    But it's come to my attention that cryovials also come in an internally threaded form, which looks like this:

    I read that the internally threaded cryotubes are supposed to provide a better seal than the externally threaded cryotubes.

    I'm trying to prevent "freezer burn" or ice crystal formation within the tube. Does having internal thread vs. external thread style cryotubes make a big difference in how well the samples are preserved at -80C over months?

    Iɽ appreciate your thoughts. Thanks and have a good day!

    When I was working with mammalian cells, we only used internally threaded. I don't, however, know of why, or when one would be favored over the other - sorry#

    Externally threaded carry about 15-20% more volume for the same general size vial.

    If you're looking to prevent freezer burn, I think it's more important that they have an O-ring

    Yes, this is the biggest factor. The internal/external threads can make a different in cap size and fitting in a container, but the o-ring is what it's all about.

    i feel like this is one of those things i've used so many times but never really questioned or learned how/why it works. what exactly do cryovials and the O ring specifically do that standard snap cap tubes dont?

    We store our single-use glycerol stocks in Matrix tubes ( which are practical because they let you store them in 96 tube racks instead of boxes and you can open 8 at a time. They've got internal threads and an O-ring. Not sure whether theyɽ be better for your application, but they've worked a lot better for us.

    I very much doubt you would see any real difference between the external vs. internal threads. Either type will create a seal that is more then adequate to preserve your samples and prevent ice crystal formation inside the tube. That being said the best way to prevent ɿreezer burn' inside your sample tubes is to let the whole vial reach room temp before opening the sample and exposing it to ambient air and the moisture in it (cold sample = condensation inside your vial.) So it has a lot more to do with sample handling then the caps ability to seal. Also as far as caps go the best seal is from push-caps or threaded caps with an O-ring. My favorite are the Micronic push-caps

    Honestly, we don't bother storing soil samples in Cryovials at all, regular 2ml screw tops work fine in the -80. Its also useful for late during RNA/DNA extraction in that those tubes with for bead beating.

    A month or two down the line? Negligible effects.

    As someone else said, you can probably get away with using regular screw top vials without much issues and you'll probably save a lot of money. I do this whenever I have a ton of cell culture samples to freeze down and when I am just using -80. As far as I know, the main difference with official cryovials is they are a bit more sturdy so when using them in liquid nitrogen, the rapid pressure change when thawing the vial will prevent cracks/explosions. In practice I'm not sure this is so much of an issue if you store your vials in the vapor phase of liquid nitrogen, but I still use nice vials even then just to be safe. I've used probably 300 standard screw-top vials in the -80 though without any signs of issue or failure. We even use standard eppendorf tubes in our -80 for some certain aliquots and have never had an issue with the vials crack or exploding.

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    This study was conducted to optimize conditions for the preparation of frozen mycoplasma reference materials suitable for evaluating alternative NAT-based mycoplasma detection methods and their comparison with conventional culture-based methods currently recommended for mycoplasma testing by regulatory agencies. Comparison of such methods, which rely on measuring naturally different biological features (i.e. GC vs CFU) requires specially prepared and well-characterized reference materials. As NAT-based methods detect mycoplasmal DNA in a sample, regardless of cell viability, the results of any comparison depend, to a great extent, on the GC/CFU ratio in the reference material used. The GC/CFU ratio is an important biological parameter that reflects the viability and aggregation of cells in a culture, both of which should be accurately assessed before selecting a reference stock to compare methods, especially, when the stock has been frozen (Dabrazhynetskaya et al. 2011 Volokhov et al. 2011 ). As many bacteria, including mycoplasmas, have a tendency to aggregate (Maniloff and Morowitz 1972 Rottem 2003 Spiglazov et al. 2004 Voloshin and Kaprel'iants 2005 ), the monitoring of cell aggregation and application of appropriate approaches to reduce aggregation are important for the proper preparation of reference materials suitable to compare methods.

    In this study, we found that aggregation of mycoplasma cells varied significantly from strain to strain, even within a single mycoplasma species. Thus, A. laidlawii strain PG8 formed large and stable cell clumps during growth in a conventional mycoplasma broth, whereas cultures of A. laidlawii strain Laidlaw A were significantly less aggregated and more dispersed when grown under the same conditions (Fig. 1). As result, the GC/CFU ratios of PG8 and Laidlaw A cultures grown for 24 h differed significantly (58 and 12, respectively Tables 2 and 3). Both visual examination of the morphology of colonies formed on surfaces of agar plates and fluorescence microscopy of stained living cells confirmed the presence of large cell aggregates in PG8 cultures, while Laidlaw A cultures showed much less cell aggregation (Fig. 1). The cell aggregates formed in PG8 cultures were stable after vigorous vortexing however, the same aggregates could be efficiently disrupted by sonication without any loss of cell viability (Table 3). As result, the GC/CFU ratio in a PG8 culture dropped from 58 to 1 after sonication, indicating that the cells became monodispersed. Therefore, sonicated cultures of those strains having an innate propensity to aggregate may yield highly viable and monodispersed cell cultures well suited to serve as reference materials for comparison of mycoplasma detection methods.

    The lack of cell walls and the specific lipid composition of the cell membranes make mycoplasma cells very susceptible to damage by freezing (McElhaney et al. 1973 Rottem et al. 1973 Raccach et al. 1975 Mazur 1984 McElhaney 1984 ). The use of optimal cooling rates and CPAs helps to establish a proper osmotic balance inside mycoplasma cells and to minimize the detrimental effects of freezing (Mazur et al. 1984 Hubalek 2003 ). We evaluated the survival of cells in cultures of A. laidlawii, Myc. gallisepticum and Myc. arginini frozen under different cooling rates in the presence or absence of different CPAs. In general, all tested mycoplasmas demonstrated the dependence of cell viability on the cooling rate. Thus, depending on mycoplasma species, an increase in the cooling rate from slow (1°C min −1 ) to rapid snapshot (60°C min −1 ) resulted in a 5-fold increase in cell survival (Fig. 2). In contrast to A. laidlawii, both Myc. gallisepticum and Myc. arginini stocks were more susceptible to the damage caused by freezing (Fig. 2). This observation can be explained by a specific lipid composition of the A. laidlawii cell membrane (Raccach et al. 1975 Lazarev et al. 2011 ) that increases the permeability of the cells and results in better survival (McElhaney et al. 1973 Mazur et al. 1984 ).

    We observed 10% DMSO and 15% glycerol as CPAs that improved the survival of all mycoplasmas tested (Fig. 2a–c). Although the choice of an optimal CPA to preserve mycoplasma cell cultures seems to depend on innate features of individual species (or even strains), both 10% DMSO and 15% glycerol are satisfactory alternatives for efficient cryopreservation. Surprisingly, 30% glycerol had a toxic effect on mycoplasma cells and, thus, cannot be recommended for cryopreservation of mycoplasma cultures. The periodic monitoring of cell viability for 1 year after freezing confirmed the high stability of all mycoplasma stocks protected with either 10% DMSO or 15% glycerol (Fig. 3a–c). Therefore, addition of CPA at an optimum concentration as well as the use of optimal cooling rates may significantly improve the survival of mycoplasma cells during the preparation and cryopreservation of reference stocks suitable to compare NAT-and culture-based methods.

    It is noteworthy that the use of proper warming rates to thaw frozen mycoplasma stocks may also increase cell recovery. Previously, it was demonstrated that thawing frozen mycoplasma cell cultures in air at ambient temperature gave an optimal warming rate that ensured efficient cell recovery (Raccach et al. 1975 Biddle et al. 2004 Boonyayatra et al. 2010 ). Some frozen mycoplasmas require special conditions for efficient recovery. Our results showed that the efficient recovery of frozen Myc. arginini G230 stocks required using anaerobic conditions for incubating agar plates that allowed us to achieve c. 99·5% efficient recovery of cells. Surprisingly, the use of aerobic conditions for agar plate incubation resulted in a four-log titre loss of Myc. arginini G230 (Fig. 4). The molecular mechanisms of this interesting phenomenon remain unknown. However, as do many other pathogens, Myc. arginini G230 utilizes l -arginine via the arginine deiminase pathway supported by anaerobic metabolism (Broman et al. 1978 Zuniga et al. 2002 Surken et al. 2008 ). The negative effect of aerobic incubation conditions we observed could be caused by inhibition of ornithine carbamoyltransferase, key enzyme of the arginine deiminase pathway, by molecular oxygen (Broman et al. 1978 ). However, we also cannot exclude the possibility of catabolic repression, as arginine-requiring Myc. arginini G230 can utilize glucose in an alternative nonfermentative way (Maniloff 1992 Pollack et al. 1997 ).